The neuroprotective role of microglial cells against amyloid beta-mediated toxicity in
organotypic hippocampal slice cultures
Richter, Maren; Vidovic, Natascha; Biber, Knut; Dolga, Amalia; Culmsee, Carsten; Dodel,
Richard
Published in:
BRAIN PATHOLOGY
DOI:
10.1111/bpa.12807
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Richter, M., Vidovic, N., Biber, K., Dolga, A., Culmsee, C., & Dodel, R. (2020). The neuroprotective role of
microglial cells against amyloid beta-mediated toxicity in organotypic hippocampal slice cultures. BRAIN
PATHOLOGY, 30(3), 589-602. https://doi.org/10.1111/bpa.12807
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Brain Pathology 0 (2020) 1–14 1
R E S E A R C H A R T I C L E
The neuroprotective role of microglial cells against amyloid
beta-mediated toxicity in organotypic hippocampal slice
cultures
Maren Richter
1,2*; Natascha Vidovic
1,3*
; Knut Biber
4,5; Amalia Dolga
2,6; Carsten Culmsee
2;
Richard Dodel
1,3*
1 Department of Neurology, Philipps-University Marburg, Marburg, Germany.
2 Institute for Pharmacology and Clinical Pharmacy, Philipps-University Marburg, Marburg, Germany. 3 Chair of Geriatric Medicine, University Duisburg-Essen, Essen, Germany.
4 Molecular Psychiatry, Psychiatric Hospital, University of Freiburg, Freiburg, Germany.
5 Department of Neuroscience, University Medical Center Groningen, University of Groningen, Groningen, the Netherlands.
6 Department of Molecular Pharmacology, Groningen Research Institute of Pharmacy, Faculty of Science and Engineering, Groningen, the Netherlands.
Keywords
Alzheimer’s disease (AD), amyloid beta (Aβ), ER stress, microglia, organotypic
hippocampal slice cultures.
Corresponding author:
Richard Dodel, MD, Chair of Geriatric Medicine, University Duisburg-Essen, Germaniastrasse 1–3, 45356 Essen, Germany (E-mail: richard.dodel@uk-essen. de)
Received 10 July 2019 Accepted 18 November 2019 Published Online
Article Accepted 26 November 2019 *Authors contributed equally to this work doi:10.1111/bpa.12807
Abstract
During Alzheimer’s disease (AD) progression, microglial cells play complex roles and have potentially detrimental as well as beneficial effects. The use of appropriate model systems is essential for characterizing and understanding the roles of microglia in AD pathology. Here, we used organotypic hippocampal slice cultures (OHSCs) to investigate the impact of microglia on amyloid beta (Aβ)-mediated toxicity. Neu-rons in OHSCs containing microglia were not vulnerable to cell death after 7 days of repeated treatment with Aβ1-42 oligomer-enriched preparations. However, when
clodronate was used to remove microglia, treatment with Aβ1-42 resulted in significant
neuronal death. Further investigations indicated signs of endoplasmic reticulum stress and caspase activation after Aβ1-42 challenge only when microglia were absent.
In-terestingly, microglia provided protection without displaying any classic signs of activation, such as an amoeboid morphology or the release of pro-inflammatory mediators (e.g., IL-6, TNF-α, NO). Furthermore, depleting microglia or inhibiting microglial uptake mechanisms resulted in significant more Aβ deposition compared to that observed in OHSCs containing functional microglia, suggesting that microglia efficiently cleared Aβ. Because inhibiting microglial uptake increased neuronal cell death, the ability of microglia to engulf Aβ is thought to contribute to its protective properties. Our study argues for a beneficial role of functional ramified microglia whereby they act against the accumulation of neurotoxic forms of Aβ and support neuronal resilience in an in situ model of AD pathology.
INTRODUCTION
Existing data strongly suggest that inflammatory processes contribute to Alzheimer’s disease (AD) pathology (36). However, microglial cells, the innate immune cells of the brain, seem to play complex roles in the progression of AD, wherein they may exert both neurotoxic and neuroprotective properties (14). In vitro data indicate that fibrils and soluble Aβ oligomers bind to various microglial surface receptors, resulting in a pro-inflammatory response of the cells (58). Pro-inflammatory microglia are thought to have a direct neurotoxic effect by releasing detrimental compounds, includ-ing reactive oxygen species (ROS) and nitric oxide (NO) (61). Recent data indicate that microglia activation patterns are highly variable and that the pro-inflammatory activity is only one possible response phenotype of microglia (21, 30, 50). In post mortem analyses of AD patients’ brains,
microglia were found to surround amyloid beta (Aβ) plaques (34, 77). In addition, in vitro studies have demonstrated that microglia engulf Aβ fibrils via receptor-mediated phagocytosis and degrade soluble oligomers either through pinocytosis (64) or by secreting Aβ-degrading enzymes, such as neprilysin and insulin-degrading enzyme (58). Nevertheless, it remains unknown whether microglial uptake of Aβ affects neurode-generation in AD pathology. Besides, accumulation of mis-folded proteins inside the ER lumen and the formation of toxic products, including ROS, are supposed to result in activation of stress responses in the endoplasmic reticulum (ER stress) (10). In AD, several human studies provide evi-dence for an involvement of ER stress in disease progression (39, 40, 59, 75). Various ER stress-related proteins have been shown to be upregulated in AD patients’ brains. The increased levels of folding proteins like 78 kDa glucose regulating protein (GRP78) and ER stress-specific caspase cleavage were
also found in AD post mortem brains (70). Organotypic hippocampal slice cultures (OHSCs) are regarded as an excel-lent model system for investigating glia–neuron interplay in the context of AD-related neurodegeneration because such cultures preserve in vivo morphology and activity of all cell types present in the hippocampal region (45, 88). Microglia in general can appear in different morphological shapes with distinct functions. These activation states are said to be reversible (67) making them highly adaptive and able to react to environmental changes. Under healthy conditions, microglial cells display a “ramified” morphology and func-tion as the brain’s controlling unit (17, 73). Thereby they are not “inactive.” Ceaselessly, with their long processes they are scanning the surrounding for invaders and pathogens or aggregated misfolded proteins (35, 49, 72). In conditions of inflammation or damage, they adapt an activation state for-merly referred to as M1 or M2, respectively (1, 73), but in 2015, Schultze and colleagues proposed a more diverse acti-vation pattern of microglia which can react to various “input signals” as it is not all black and white (76). In contrast to pure, isolated microglial cultures that display various morphologies ranging from needle-like to amoeboid in shape, in OHSCs, microglia maintain their natural environment and therefore show a predominately ramified morphology (33, 88). In addition, neurons in slice cultures have intact synapses with an in vivo-like distribution of receptors (e.g., NMDA receptors) that results in drug sensitivity and EC50 values
similar to those observed in living organisms (38). OHSCs are therefore an ideal tool to investigate the function of microglia in an in vivo-like cellular environment (33, 65).
In the present study, we used OHSCs to examine the impact of microglial cells on Aβ-mediated effects. Neuronal death induced by treatment with Aβ, and the mechanisms underlying were characterized in the presence or absence of microglia.
MATERIALS AND METHODS
Preparation and cultivation of OHSCs
The preparation of organotypic hippocampal slice cultures was performed as previously described (79), with minor modifications. Postnatal day 1–4 C57Bl/6J mice (Janvier, Le Genest-saint-Isle, France or Charles River Laboratories, Sulzfeld, Germany) were sacrificed by decapitation, and their brains were dissected. Hippocampi were dissected, cut into 375-µm thick slices using a McIlwain tissue chop-per (Mickle laboratory engineering, Surrey, UK) and cul-tivated on membrane inserts (Millicell cell culture inserts, 0.4 µm pore size, Merck Millipore, Darmstadt, Germany) in 6-well plates. Per well, six slices were arranged on a membrane insert, with each well containing 1.2 mL sterile-filtered (Whatman syringe filters, 0.2 µm pore size PES, Merck, Darmstadt, Germany) culture medium (for 100 mL: 41.6 mL aqua ad injectabilia, 5 mL Minimum Essential Medium 10×, 25 mL Basal Eagle’s + Earle’s Medium, 25 mL HyClone Donor Equine Serum, 1 mL GlutaMax, 1.44 mL 45% glucose solution, pH 7.2; all life
technologies, Carlsbad, USA), The hippocampal slices were incubated in a humidified atmosphere containing 5% CO2
at 35°C.
To specifically deplete microglia, 100 µg/mL clodronate disodium salt (Merck Millipore, Darmstadt, Germany) was added to the medium, as previously described (54). The day after preparation, slices were washed with 1× PBS (Merck Millipore, Darmstadt, Germany), and inserts were transferred into a new 6-well plate containing fresh culture medium without clodronate. Culture medium was changed every 2–3 days, and treatment started 6–7 days after preparation. Amyloid beta (Aβ1-42) application to OHSCs
The preparation of Aβ1-42 oligomer-enriched solutions was
performed as previously described (23). A total of 1 mg Aβ1-42 (Bachem, Bubendorf, Switzerland) were dissolved in
10% ammonia (Sigma-Aldrich, Taufkirchen, Germany) to a final concentration of 1 mg/mL and then aliquoted into low-binding tubes (Eppendorf, Hamburg, Germany). Afterward, the aliquots were freeze-dried to obtain 25 µg protein and stored at −80°C until use. Before used in OHSCs, Aβ1-42 aliquots were dissolved in aqua ad injectabilia
(1 mg/mL) and filled up with 100 mM Tris, 50 mM NaCl (Sigma-Aldrich, Taufkirchen, Germany), pH 7.4, to obtain a 58 µM solution. As a negative control, the described Tris/NaCl buffer was prepared without adding Aβ1-42 peptide.
The Aβ1-42 solution and the solution containing the buffer
without the peptide were each stirred for 48 h at 1400 rpm at room temperature. For the treatment of OHSCs, 2 µL of the freshly prepared oligomer-enriched 58 µM Aβ1-42
solution was dropped on top of each of the slices, con-sidering six slices per well containing 1.2 mL culture medium; the final concentration of Aβ1-42 is 0.6 µM. To obtain higher
concentrations, oligomeric Aβ1-42 solution additionally was
applied on top of the slices at a concentration of 1 µM (3 µL per slice), to obtain a final concentration of 1.6 µM in the well. It is not recommended to add more than 3 µL of solution at once on top of the slices because the excess liquid will slide off and end up in the media too quickly. To obtain lower (0.3 µM) concentrations of Aβ1-42, the
stock solution (58 µM) was diluted in culture medium to 30 µM, and then, 2 µL of this dilution was dropped on top of the slices to obtain a final concentration of 0.3 µM in the well. By dropping Aβ1-42 directly on top of the
slices, it is more likely to be taken up and interacts with the cells than by adding it to the media. The Aβ1-42 peptide
was re-applied on top of the slices with every media change, which occurred 3 times over one week.
Detection of neuronal cell death
Propidium iodide (PI) staining was performed with immu-nostaining to detect neuronal cell death in OHSCs. Following Aβ1-42 treatment, PI was added to the medium
at a final concentration of 10 µg/mL, and the cultures were incubated for 1 h at 35°C. Afterward, the dye was removed, and the OHSCs were washed and fixated in 4% paraformaldehyde (PFA) solution. After several washes,
immunostaining was performed for neurons (NeuN) and microglial cells (Iba1). ImageJ (Rasband, W.S., ImageJ, U. S. National Institutes of Health, Bethesda, MD, USA) was used to analyze neuronal cell death in OHSCs. An oval selection tool was used for the hippocampal regions (CA1+3 and DG). The threshold was adjusted for far-red (Iba-1), red (PI) and green (NeuN). The area covered with far-red (Iba-1), red (PI) and green (NeuN) within the selected oval area was measured. Only slices with less than 10% of the hippocampus regions CA1+3 and DG covered with Iba1-positive microglia were used for quan-tification of cell death. Therefore, the microglial presence was calculated by dividing the magenta area (Iba-1) by the green area (NeuN). Neuronal cell death within one hippocampal slice was calculated as the red area (PI) divided by the green area (NeuN). The presented data were pooled from several hippocampal slice cultures rep-resenting three independent experiments per condition. Detection of activated caspases
CaspACE™ FITC-VAD-FMK (Promega, Madison, USA) was
used to detect activated caspases inside the cells. The OHSC culture medium was removed, and medium containing CaspACE™ FITC-VAD-FMK at a final concentration of 5 µM
was applied with the Aβ1-42 treatment. The cultures were then
incubated for 48 h. At the end of the treatment period, the slices were washed, fixed with 4% PFA and immunostained.
ImageJ was used to analyze caspase activation in OHSCs. The oval selection tool was used so select hippocampal regions (CA1+3 and DG). The threshold was adjusted for green (CaspACE™ FITC) and far-red (NeuN), and the areas
express-ing each color within the selected oval area were measured. Neuronal caspase activity within one hippocampal slice was calculated as the green area divided by the far-red area. Thioflavin T staining
After respective treatments, the OHSC culture medium was removed from each culture, and the slices were washed in 1× PBS (Merck Millipore, Darmstadt, Germany) and fixed with 4% PFA at 4°C. To stain amyloid deposits, Thioflavin T (ThioT, Sigma-Aldrich, Taufkirchen, Germany) was diluted to a final concentration of 0.1% in aqua bidest. A ThioT working solution was mixed 1:1 in 100% ethanol and applied to slices for 10 minutes at RT in the dark. Slices were then washed with 80% ethanol and aqua bidest, and the procedure for immunostaining was then performed.
ImageJ was used to analyze ThioT staining in OHSCs. The oval selection tool was used to select hippocampal regions (CA1+3 and DG). After adjustment for the appro-priate threshold value, the amount of ThioT within the selected oval area was measured.
Immunocytochemistry
The PFA-fixed slices were cut off from the membrane and transferred into 3% BSA solution for blocking. Afterward, the slices were incubated with primary antibodies (anti-NeuN,
Merck Millipore, Darmstadt, Germany and anti-Iba1, Wako Chemicals GmbH, Neuss, Germany) overnight at 4°C. After the tissues were washed three times for 20 minutes, fluo-rescently labeled secondary antibodies (all Invitrogen, Karlsruhe, Germany; anti-mouse IgG (H+L) AlexaFluor488, anti-mouse IgG (H+L) AlexaFluor 546, anti-rabbit IgG (H+L) AlexaFluor647, and anti-rabbit IgG (H+L) AlexaFluor546, all 1:1000) were applied, and the slices were incubated for 2 h at room temperature. Again, after the slices were washed three times for 20 minutes each, the slices were transferred onto object slides and covered with fluorescence mounting medium (Dako, Santa Clara, USA), and a coverslip was placed on top.
Analysis of the OHSC staining
All OHSC staining experiments were analyzed using confocal laser scanning microscopy with a LSM 510 META Laser Scanning Microscope (Carl Zeiss, Jena, Germany). Light was collected through a Plan-Neofluor 10×/0.3 objective to obtain overview pictures of the hippocampal slices or a 63×/1.4 Oil DIC objective for single-cell analysis. 3D single-cell reconstruc-tions were performed using z-stacks with 0.4 μm image intervals. For the Alexa Fluor 488 antibody, CaspACE FITC-VAD-FMK and ThioT staining, the fluorescence was excited at 488 nm, and emissions were detected between 505 and 550 nm. For the Alexa Fluor 546 antibody and PI staining, fluorescence was excited at 543 nm, and emissions were detected using a 560 nm long pass filter. For the Alexa Fluor 647 antibody, fluorescence was excited at 488 nm, and emissions were detected between 680 and 800 nm.
Western blot analysis
Protein concentrations were determined using a Pierce BCA Protein Assay kit (Thermo scientific, Rockford, USA) according to the manufacturer’s protocol. XCell Sure Lock Mini-Cell Electrophoresis System was used to separate the proteins (Invitrogen, Karlsruhe, Germany). Protein samples were mixed with NuPAGE® LDS Sample Buffer (4×) (Life technologies, Carlsbad, USA) supplemented with 0.05 M dithiotreitol (Sigma-Aldrich, Taufkirchen, Germany) and then loaded into 10 1.5-mm NuPAGENovex 4%–12% Bis-Tris Protein Gels wells together with SeeBlue Plus2 Pre-stained Protein Standard (both Life technologies, Carlsbad, USA). Blots were then transferred onto nitrocellulose membranes (Whatman, Maidstone, UK) using an XCell II Blot Module (Invitrogen, Karlsruhe, Germany), and the membranes were reversibly stained using a MemCode Reversible Protein Stain Kit (Perbio Science, Bonn, Germany) to visualize that the protein had transferred. After blocking the membranes against nonspecific binding, they were incubated with primary antibodies (anti-Cas-pase-12, Cell Signaling, Danvers, USA; anti-GRP78, Santa Cruz, Dallas, USA; anti-Tubulin, Sigma-Aldrich, Taufkirchen, Germany; or anti-Vinculin, Sigma-Aldrich, Taufkirchen, Germany) overnight and then with conjugated secondary antibodies (anti-mouse peroxidase-conjugated; anti-rabbit peroxidase-peroxidase-conjugated; or anti-goat
peroxidase-conjugated, Vector Laboratories, Peterborough, UK) for 1 hour. To visualize bands, the membranes were incubated with SuperSignal West Dura Extended Duration Substrate (Thermo Scientific, Rockford, USA) and exposed to an autoradiographic film (CL-Xposure Film, Thermo Scientific, Rockford, USA). Films were scanned using a GS-800 Calibrated Densitometer, and densitometric analysis was performed using Quantity One software (both Bio-Rad Laboratories, Munich, Germany). Quantification was car-ried out by calculating the fold increase of treatment conditions in comparison to the control condition. Cytokine ELISA for IL-6 and TNF-α
For the IL-6 and TNF-α ELISAs, mouse IL-6 ELISA Ready-SET-Go! and mouse TNF-α ELISA Ready-SET-go! kits (both eBioscience, San Diego, USA), respectively, were used according to the manufacturer’s protocol. Briefly, the supernatant was collected from each OHSC during every media change (3 times), and these samples were combined before use. ELISA 96-well plates (Microlon, Greiner, Frickenhausen, Germany) were incubated overnight at 4°C with capture antibody solution in 1× coating buffer. The plate was washed with 1× PBS supplemented with 0.05% tween 20, blocked with 1× assay diluent for 1 h and then washed again. Afterward, cytokine standards and samples were applied and they were incubated for 2 h. After the plates were washed, they were incubated with a detection antibody dilution in 1× assay diluent for 1 h, followed by an additional washing step. Streptavidin-HRP was incu-bated for 30 minutes in 1× assay diluent. Before incubation of the plates in tetramethylbenzidine (TMB) substrate, they were washed 2 times. After 15 minutes, the reaction was stopped using 5% H2SO4 (Carl Roth, Karlsruhe, Germany).
Absorbance was measured at 450 nm using a plate reader (Infinite M200, Tecan, Männedorf, Switzerland).
Griess assay
Nitric oxide (NO) production was measured using a col-orimetric method based on a chemical diazotization reaction using Griess reagent. During every media change, the OHSC supernatant was collected (3 times), and they were combined prior to use. The samples and standards were transferred to a 96-well plate (Greiner, Frickenhausen, Germany) and incubated for 10 minutes with reagent I (1% (m/V) sulfa-nilamide, 5% (V/V) in phosphoric acid) at room temperature in the dark. Afterward, reagent II (0.1% (V/V) naphthyl-ethylenediamine dihydrochloride) was added, and the plates were incubated for 10 minutes in the dark. Absorbance was measured using a plate reader (Infinite M200, Tecan, Männedorf, Switzerland) at 550 nm.
Preparation and cultivation of isolated microglia
Adult mice (Janvier, Le Genest-saint-Isle, France) were sac-rificed via cervical dislocation. Embryonic mice (E13.5) were decapitated and brains were dissected. Microglial cells were
obtained from the embryonic cortex and collected into Leibovitz L-15 medium (PAA Laboratories, Pasching, Austria). Afterward, the cells were homogenized and centrifuged at 300 × g for 5 minutes; the remaining pellet was re-suspended in microglia culture medium (DMEM/F-12, Fetal Calf Serum, 100 U/mL penicillin and 100 µg/mL streptomycin and 200 mM
l-glutamine, pH 7.2; all PAA Laboratories, Pasching, Austria).
The cells were cultured in polyethylenimine-coated 6-well plates. The next day, half of the medium was changed. On day 7, the medium was supplemented with 5 ng/mL granu-locyte–macrophage colony-stimulating factor (Roche, Basel, Switzerland), as previously described (12). On day 13, astro-cytes were removed from the culture via trypsinization using 0.25% trypsin at 37°C. Microglial cells were re-suspended and re-plated onto PEI-coated 24- or 96-well plates at a density of 100 000–200 000 cells per mL. The cells obtained using this protocol were mainly microglia (>95%), as shown by CD11b flow cytometric analysis.
Real-time measurements of cell impedance The xCELLigence Real-Time Cell Analyzer RTCA-MP system (Roche diagnostics, Rotkreuz, Switzerland) was used to detect changes in cellular impedance that resulted from changes in microglial morphology (11). To subtract the background, the impedance of the primary microglia medium was recorded prior to seeding the cells. Afterward, 15 000 primary micro-glial cells were seeded in 100 µL cell culture medium per well in 96-well format E plates (Roche diagnostics, Rotkreuz, Switzerland). After 24 h of incubation, the cells were treated with LPS (a positive control (11)) or Aβ1-42. Cell index values
were recorded during the whole treatment period using RTCA software 1.2 (Roche Diagnostics, Rotkreuz, Switzerland). Statistical analysis
All data are shown as the mean ± standard deviation (SD). Multiple comparisons were performed using analysis of vari-ance (ANOVA) followed by Scheffé’s post hoc test. P < 0.05 was used as threshold p-value to define significance. Calculations were performed using the Winstat standard statistical software package (R. Fitch Software, Bad Krozingen, Germany).
RESULTS
Repeated treatment with oligomer-enriched preparations of Aβ1-42 for 7 days does not
induce neuronal cell death in OHSCs
OHSCs obtained from wild-type C57BL/6J mice, which main-tained neuronal and microglial cells in the in vivo formation (Figure 1A), were used as an in situ model to study Aβ1-42
-induced neurotoxicity. OHSCs were challenged with increasing concentrations of oligomer-enriched Aβ1-42 preparations (300–
1600 nM) and were subsequently double-stained with Neuronal Nuclei (NeuN) antibody and PI (Figure 1B). Neuronal survival in the dentate gyrus (DG) and cornu ammonis (CA1+3) regions was not significantly affected in OHSCs following treatment with Aβ1-42 for up to one week (Figure 1E).
Depletion of microglial cells enables Aβ1-42
-induced neuronal death in OHSCs
The induction of apoptosis by the use of the macrophage toxin clodronate is a well-established method for depleting microglial cells both in vitro (56) and in vivo (15). Treatment
of OHSCs with clodronate has been shown to selectively deplete microglial cells without affecting other cell types (45). Clodronate (0.1 mg/mL) was applied to freshly obtained OHSCs for 24 h, resulting in OHSCs without Iba1-positive microglial cells and unaffected neuronal formations (Figure 1C). To
investigate the role of microglia in Aβ1-42-induced effects,
microglia-depleted OHSCs were incubated with increasing concentrations of oligomer-enriched preparations of Aβ1-42
(300–1600 nM) and then analyzed to determine whether there were changes in neuronal cell death (Figure 1D). Concentrations starting from 600 nM Aβ1-42 significantly increased neuronal
cell death in OHSCs that had been depleted of microglia compared to untreated control OHSCs (Figure 1F).
Microglial cells prevent ER stress and caspase cleavage in Aβ1-42-challenged OHSCs
Aβ1-42 oligomers induced neuronal apoptosis via
caspase-dependent pathways (2, 70). To investigate neuronal caspase activity in OHSCs, we applied a fluorescent analog of the pan-caspase inhibitor Z-VAD-FMK, which allows for detec-tion of activated caspases, over a time period of 48 h and then performed counterstainings for NeuN. We did not detect a significant increase in neuronal caspase cleavage after Aβ1-42
treatment within 48 h. However, the depletion of microglial cells strongly enhanced the expression of activated caspases after the challenge with Aβ1-42 (Figure 2A,B). In addition,
previous studies have indicated that treatment with Aβ potenti-ates ER stress-induced expression of caspase-12 in OHSCs (43). According to recent investigations, Aβ oligomers are able to move between the intracellular and the extracellular space (22, 82) and may mediate toxicity on both levels. Interestingly, Aβ oligomers were detected in the ER of hippocampal neu-rons in a mouse model of AD where they are supposed to induce organelle functional impairment resulting ultimately in cell death (86). Furthermore, recent evidence suggests that Aβ oligomers induce defects in lipid bilayers, thus, inducing rapid calcium influx (48) which may affect the ER as the main intracellular calcium store. Therefore, we next aimed to investigate whether Aβ-mediated toxicity in microglia-depleted OHSC may be accompanied by upregulation of ER stress marker proteins. We found that cleavage of caspase-12 occurred only in OHSCs that had been depleted of microglial cells and was enhanced by Aβ1-42 treatment, as shown by Western
Blot analysis (Figure 2C). We next investigated a second ER stress marker, the ER resident chaperone GRP78. GRP78 protein expression was not affected by either microglial cell depletion or treatment alone. However, Aβ1-42 treatment in
microglia-depleted OHSCs resulted in enhanced GRP78
expression (Figure 2D), suggesting that microglial cells inhibited Aβ1-42-induced ER stress.
Microglial cells provide their protective effect in a ramified morphological state
Under physiological conditions, microglial cells display a rami-fied morphology that has long been defined as their “resting phenotype.” Nevertheless, recent data indicate that ramified microglia are not “resting” but are actively scanning their environment to maintain tissue homeostasis (21, 30, 50).
We treated OHSCs with Aβ1-42 for 7 days and then
inves-tigated the morphology of microglia using Iba1 staining. LPS treatment for 24 h, which was used as a positive control to induce full microglial pro-inflammatory activation, is characterized by a morphological transition toward amoeboid cells. It is described here that Aβ1-42 treatment did not change
the morphological properties of microglia compared to microglia from untreated OHSCs (Figure 3A). For an addi-tional quantitative assessment of microglial responses, we used real-time impedance measurements of pure microglia cultures. These measurements provide information related to changes in microglial morphology, which were continu-ously monitored for the entire Aβ1-42 challenge period. LPS
induced marked alterations in microglial structural features, as detected using xCELLigence measurements. Treatment with high concentrations (20 µM) of oligomer-enriched Aβ1-42
preparations, however, did not induce changes in microglial morphology, as measured using an xCELLigence system (Figure 3B).
In addition to morphological changes, the pro-inflam-matory activation of microglia resulted in the increased production of pro-inflammatory cytokines and NO release. Therefore, levels of pro-inflammatory cytokines were exam-ined after 7 days of Aβ1-42 treatment. LPS treatment for
24 h was used as a positive control. In OHSCs in which microglial cells were present, Aβ1-42 treatment alone did
not significantly increase IL-6 (Figure 4A), TNF-α (Figure 4B) or NO levels (Figure 4C). According to these data, Aβ1-42 treatment did not result in microglia with a
pro-inflammatory phenotype. Therefore, a morphological transi-tion or pro-inflammatory activity of ramified microglia seemed to be unnecessary for improving neuronal survival after Aβ1-42 treatment in OHSCs.
Figure 1. Selective depletion of microglial cells by clodronate facilitates Aβ1-42-induced neurotoxicity in OHSCs. A. Representative image of an untreated OHSC stained with PI (dead cells), NeuN (neurons) and Iba1 (microglia) after 13 days in culture. B. Representative image of an OHSC after 7 days of repeated treatment with 600 nM Aβ1-42. The OHSC was
stained with PI, NeuN and Iba1. C. Representative image of an untreated OHSC depleted of microglia by treatment with clodronate (100 µg/mL, 24 h) and then stained with PI (dead cells), NeuN (neurons) and Iba1 (microglia) after 13 days in culture. D. Representative image of a microglia-depleted OHSC after 7 days of repeated treatment with 600 nM Aβ1-42. The OHSC was stained with PI, NeuN and Iba1. E.
Quantification of neuronal cell death after 7 days of treatment with different concentrations Aβ1-42. OHSCs were stained with PI and NeuN,
and the amount of dead neurons [%] was calculated as the red area (PI)
divided by the green area (NeuN) in the hippocampal regions CA1+3 and DG (mean ± SD; 11–19 individual slices per condition from 3 independent preparations, n.s. P > 0.05). F. Quantification of neuronal cell death after 7 days of treatment with different concentrations of Aβ1-42 in OHSCs
depleted of microglia. OHSCs were stained with PI, NeuN and Iba1. Only slices in which less than 10% of the hippocampal regions CA1+3 and DG were covered by Iba1-positive microglia were used for quantification. The amount of dead neurons [%] was calculated as the red area (PI) divided by the green area (NeuN) in the hippocampal regions CA1+3 and DG (mean ± SD; 7–15 individual slices per condition from 3 independent preparations, n.s. P > 0.05, **P < 0.01, ***P < 0.001 compared to the untreated control).
Phagocytosis of Aβ1-42 contributes to the
neuroprotective effects of microglia
Previous in vitro studies revealed the ability of microglia to take up Aβ (18, 20, 29, 64). However, it is unknown whether microglial uptake of Aβ reduces peptide toxicity to contribute to neuronal survival. Here, we performed Thioflavin T (ThioT) staining to investigate whether the increase in neuronal death observed after Aβ1-42 treatment in microglia-free OHSCs was
accompanied by an enhanced Aβ plaque load. First, we
confirmed that microglial cells were able to engulf ThioT-positive Aβ1-42 in OHSCs using z-stack analysis of ThioT and
Iba1 expression in single microglial cells (Figure 5A). Next, we compared ThioT staining in OHSCs with and without microglial cells (Figure 5B,C). The analysis of ThioT-positive areas of microglia-depleted OHSCs following Aβ1-42 treatment
for 48 h did not result in a significant difference compared to non-treated OHSCs (Figure 5C). In contrast, significant increases in hippocampal areas positive for ThioT were observed
Figure 2. Aβ1-42 treatment in OHSCs results in ER stress and caspase cleavage only in the absence of microglial cells. A. Representative images from different treatment conditions after CaspACE™
FITC-VAD-FMK staining for activated caspases and NeuN immunostaining for neurons. B. Quantification of neuronal caspase activity in OHSCs after 48 h of treatment with 600 nM Aβ1-42. The caspase staining within one
hippocampal slice was calculated as the green area (CaspACE FITC-VAD-FMK positive) divided by the far-red area (NeuN) (mean ± SD; 8–17 individual slices per condition from 3 independent preparations,
***P < 0.001 compared to treatment for 48 h with Aβ1-42 with microglial
cells present). C. Representative Western blot for caspase-12 and cleaved caspase-12 after 48 h of treatment with 600 nM Aβ1-42 with and
without microglial cells present. Corresponding densitometric quantification is shown (mean ± SD; n = 3). D) Representative Western blot for GRP78 after 48 h of treatment with 600 nM Aβ1-42 with and
without microglial cells present. Corresponding densitometric quantification is shown (mean ± SD; n = 3).
after 7 days of treatment with Aβ1-42 in microglia-free slices,
indicating that Aβ plaques formed in this ex vivo model. To further investigate the impact of microglial Aβ1-42 uptake
on neuronal survival in OHSCs, we applied cytochalasin D, a blocker of actin polymerization that inhibits the phagocytic activity of microglia (64, 74) at a concentration that was barely neurotoxic in OHSC (52). Treatment with cytochalasin D resulted in a significant increase in the hippocampal area that was positive for ThioT, suggesting efficient inhibition of microglial Aβ1-42 intake (Figure 5D). Inhibiting microglial uptake
mechanisms with cytochalasin D increased the number of dead neurons that were positive for PI staining following treatment with Aβ1-42 (Figure 5E), indicating that Aβ1-42
phago-cytosis contributes to microglia-mediated protective effects.
DISCUSSION
Our study provides evidence that the presence of ramified microglia reduces Aβ1-42-mediated neurotoxicity in postnatal
wild-type OHSCs. This reduction in neuronal cell death was accompanied by a reduced Aβ load, suggesting an important neuroprotective role for microglial phagocytosis.
OHSC as a model to investigate neuron–glia interactions
To characterize and understand the roles of microglia in AD pathology, it is essential to use appropriate models. For many years, pure, isolated microglial cultures were used
for investigation. However, one of the great disadvantages of the use of isolated microglial cultures is the lack of inhibitory signals that arise from neurons (7, 32, 37). Accordingly, several expression studies have provided evidence that standard cultured microglia are more similar to periph-eral myeloid cells than to in vivo microglia (4, 6). This may explain why most of the studies that investigated the influ-ence of isolated microglia on neuronal survival described the neurotoxic potential of these cells (5, 24, 47), which is in contrast to our current findings. In the present study, OHSCs were used as a model system to investigate the interplay between microglia and neurons in a pathology relevant to AD. Because microglia in OHSCs maintain their
in vivo environment and exhibit a ramified morphology Figure 3. Microglial morphology after Aβ1-42 treatment. A. Microglial
morphology was assessed using Iba1 staining in untreated OHSCs and after 24 h of treatment with LPS 1 µg/mL or 7 days of treatment with Aβ1-42 600 nM. B. Real-time detection of morphological changes in
untreated pure microglial cultures and after treatment with 1 µg/mL LPS or 20 µM Aβ1-42. Values were normalized to 1 before treatment
(mean ± SD; n = 6).
Figure 4. Microglial release of pro-inflammatory mediators after Aβ1-42 treatment. A-C. The release of pro-inflammatory mediators was investigated in OHSC supernatants using ELISA measurements of TNF-α and IL-6 levels and Griess assays for NO release. OHSCs were treated with 100 µg/mL LPS for 24 h or 600 nM Aβ1-42 for 7 days
(78, 88) that is comparable to their appearance in vivo (64), the use of OHSCs is thought to be more appropriate for disease modeling than cultures containing single-cell types. The reagent bisphosphonate clodronate has long been used
to deplete microglial cells in vitro as well as in vivo (15, 56), and it has never affected the number and integrity of neurons in these different studies. Hence, the depletion of microglial cells from OHSCs using clodronate serves as an
excellent tool for investigating the impact of microglia on Aβ-mediated effects.
Aβ1-42-mediated effects on neurons in OHSCs In the present study, exogenous Aβ1-42 peptide, in nanomolar
concentrations (up to 1.6 µM), was repetitively applied to OHSCs for one week, starting on days 6–7 in culture. The Aβ1-42 concentrations used were in the range of the
concentrations described for Aβ levels in AD patients’ brains (71). Nevertheless, treatment with Aβ1-42
oligomer-enriched preparations did not result in neuronal cell death in OHSCs, as shown by the results of PI staining (Figure 1). This is in line with previous studies that showed that applying different Aβ species (Aβ1-28, Aβ25-35, Aβ1-40) for
various durations (24 h–7 d) failed to trigger neuronal cell death in OHSCs (63, 87, 90). In contrast, numerous studies have described the neurotoxic potential of oligomer-enriched Aβ preparations in OHSCs. However, in most of these studies, OHSCs were treated in serum-free (55, 80, 89) or serum-reduced (41, 54) media, or the Aβ treatment was combined with other detrimental stimuli (3, 43) to boost toxicity. In addition, higher concentrations, ranging from 2 to 10 µM Aβ1-42, resulted in significant neuronal
cell death in OHSCs after 24–48 h of treatment (42, 51, 93). The concentrations used in these experiments are, however, thought to be far from those observed under physiological settings in AD.
Previous studies have shown that Aβ-mediated cell death is caspase-dependent in pure neuronal cultures (2, 70). Regarding OHSCs, treatment with Aβ25-35 and Aβ1-42 at high
micromolar concentrations resulted in the cleavage of cas-pase-3 (19), suggesting its involvement in Aβ-mediated apop-tosis. In our experimental setting, staining for activated caspases indicated that Aβ1-42-mediated cell death mechanisms
are caspase-dependent in the absence of microglia (Figure 2A,B). Furthermore, the enhanced expression of ER stress marker proteins implies that ER stress pathways may be involved in Aβ1-42-induced toxicity in absence of microglia
(Figure 2C,D). Aβ treatment has been previously demonstrated to result in caspase-12-mediated cell death mechanisms and enhanced expression levels of GRP78 in primary neuronal cultures (70, 94). In contrast, cleavage of caspase-12 or the upregulation of GRP78 after treatment with Aβ25-35 was not
detected in OHSCs (43). In addition, treatment with Aβ for 48 h resulted in the cleavage of caspase-12 only in old OHSCs that were cultured for 7 weeks (44), suggesting that ER
stress pathways may be involved in Aβ-mediated cell death in neurons. However, the presence of functional ramified microglia may be sufficient to block this route of toxicity. Aβ1-42-mediated effects on microglia in OHSC
As described earlier, activated microglia are found in the sur-rounding of amyloid plaques indicating an interaction with amyloid oligomers and fibrils (16). Supporting evidence for an involvement of amyloid oligomers in the activation of microglial cells is derived from the fact that on microglial cells, receptors were found capable of binding to Aβ1-42 (58, 69).
Binding to these receptors activates a signaling cascade resulting in the expression of cytokines of either pro- or anti-inflammatory origin or leads to the internalization of abeta molecules. Furthermore, when microglia face such inflammation sites for the first time, they are said to be “primed” and undergo a change in their morphology toward an amoeboid appearance (34, 60). In the present study, however, treatment of OHSCs with Aβ1-42 oligomer-enriched
preparations for 7 days did not result in the classical signs of microglial activation, including significant increases in the release of the pro-inflammatory cytokines IL-6 and TNF-α and NO secretion (Figure 4) and a switch in morphology toward an amoeboid phenotype (Figure 3). Besides, the above described activation states, Wyss-Coray and Rogers summa-rized in 2012, another phenotype which is “acquired deac-tivation” (9, 26, 92). The term was first introduced by Gordon in 2003 and differs from the “classical alternative activation state,” also formerly known as M2, in the way, that next to cytokines (Interleukins (IL-10); Tumor growth factor-β (TGF-β)) also apoptotic cells themselves can give rise to this second alternative state (9, 25, 68). Microglial cells with this phenotype are said to be “immunosuppressive” and are not capable of producing cytokines of either pro- or anti-inflammatory origin. Besides, their main function is to take up apoptotic cells. The reason we did not observe an amoe-boid phenotype in our experimental setup could be that the microglial cells were not activated classically by amyloid oligomeric structures, but still “active” in the way of clear-ing cellular debris by maintainclear-ing a ramified phenotype.
In contrast, previous studies that used the same Aβ1-42
oligomer-enriched preparations but at higher micromolar concentrations (10 µM) demonstrated IL-6 and TNF-α release from primary microglial cultures (23), thereby indicating a potential difference between microglia used in suspension cultures and microglia in ex vivo models. However, in former
Figure 5. Phagocytosis of Aβ1-42 contributes to the neuroprotective effects of microglia. A. A 3-D image was created using z-stacks showing ThioT and Iba1 staining in a single microglial cell from an OHSC that was treated with 600 nM Aβ1-42 for 7 days. Cross indicates sagittal and
transversal section planes. B. Representative images showing ThioT, NeuN and Iba1 staining in OHSCs after 7 days of repeated treatment with 600 nM Aβ1-42 with (1) and without (2) microglial cells present; a
fivefold magnification of ThioT staining is given. C. Quantification of ThioT-positive hippocampal areas after treatment with 600 nM Aβ1-42 for
the indicated times (mean ± SD; 12–18 individual slices per condition from 3 independent preparations, ***P < 0.001 compared to 7 days of
Aβ1-42 treatment in the presence of microglia). D. Assessment of ThioT
staining after treatment with Aβ1-42 in the presence and absence of the
phagocytosis blocker Cytochalasin D (mean ± SD; 8–13 individual slices per condition, **P < 0.01 compared to treatment with Aβ1-42). E.
Quantification of neuronal cell death after 7 days of treatment with Aβ1-42
and Cytochalasin D. OHSCs were stained with PI and NeuN, and the amount of dead neurons [%] was calculated as the red area (PI) divided by the green area (NeuN) in the hippocampal regions CA1+3 and DG (mean ± SD; n = 7–9 per condition, ***P < 0.001 compared to the control).
OHSCs studies, treatment with Aβ1-40 (8) and Aβ25-35 (41)
at higher micromolar concentrations resulted in the enhanced expression of IL-6 and TNF-α, while nanomolar concentra-tions of Aβ1-42 (20 nM) did not induce detectable increases
in IL-6 and TNF-α release (62), suggesting that either high concentrations or different Aβ preparations (or a combina-tion of both) are sufficient to promote the release of pro-inflammatory cytokines from microglia in OHSCs.
One previous investigation using an OHSC model dem-onstrated that stimulation with Aβ1-42 oligomers results in
enhanced NO release from microglia (62). The authors identified NO as the major mediator of Aβ1-42
oligomer-induced microglial neurotoxicity. Therefore, the absence of NO in the present study may explain the lack of neuro-toxicity induced by the Aβ1-42 oligomer preparation we used.
Nevertheless, it remains unknown why these two different studies reached such different results. Differences in the Aβ oligomerization protocols used may partly explain this phe-nomenon. In the former study by Maezawa and colleagues, however, NO-mediated microglial neurotoxicity was accom-panied by the presence of an amoeboid phenotype (62), indicating that this kind of morphologically activated micro-glia can also promote the death of neurons.
Ramified microglia provide protection against Aβ1-42 toxicity
In the present study, the presence of ramified microglia prevented Aβ1–42-mediated neuronal cell death, while the
depletion of microglial cells facilitated neuronal tissue dam-age, suggesting a protective effect by microglia.
OHSCs were obtained from postnatal, healthy, wild-type mice, microglia were uniformly distributed throughout the hippocampal slices used in the cultures (Figure 1A), and they exhibited a ramified morphology (Figure 3A), suggest-ing that the functional integrity of the microglia was pre-served in the experimental setting. Interestingly, treatment of the slices with Aβ1–42 for 7 d did not affect their ramified
morphological state (Figure 3A).
The neuroprotective properties of non-ramified microglial cells are well described in OHSC models of excitotoxicity (85) or stroke in vivo (15, 57) and to a lesser extent for ramified microglia (65, 88). Furthermore, a few studies have been performed to investigate the role of microglial cells in in vivo and ex vivo models of neurodegenerative diseases. In OHSCs obtained from CD11b-HSVTK mice, inducing the removal of microglial cells with ganciclovir increased prion titers 15-fold, suggesting a beneficial role for microglia in prion infections (13). However, in a mouse model of ALS in which the mice carried a superoxide dismutase (SOD) mutation, depleting microglial cells did not affect disease progression (27), whereas in vivo microglial depletion from wild-type CB57BL/6 mice was shown to worsen brain ischemia and to enhance neuroinflammation by upregulating important pro-inflammatory cytokines as well as increased neuronal cell death (46, 81). Regarding transgenic mouse models of AD, depleting microglial cells did not affect Aβ plaque load
in vivo (28). However, as the here presented data and recent
findings suggest that microglia prevent the formation of Aβ
plaques (33), this lack of effect on Aβ plaque load in AD models might be due to rather late time point were microglia have been depleted. It is moreover important to note that severe neurodegeneration, like that observed in AD patients, does not take place in the used transgenic animals (28), which makes it difficult to address the impact of microglia on AD-related neurodegeneration in these models in vivo. Therefore, clodronate-induced microglial depletion in OHSCs is an excellent tool for addressing this particular point. To the best of our knowledge, the present study is the first to demonstrate a beneficial effect for ramified microglia in Aβ-mediated neurotoxicity in an ex vivo model of AD.
Several mechanisms were described that may contribute to the microglia-mediated neuroprotective effects observed in models of Aβ1-42-mediated toxicity. These include the
release of neurotrophic factors that may provide trophic support to surrounding neurons and neuronal processes (21) in addition to the ability to take up toxic substances, such as glutamate, from the extracellular environment and to render them harmless (30). Here, depleting microglial cells increased the Aβ load and resulted in neuronal apoptosis. Furthermore, efficiently blocking microglial uptake mecha-nisms with cytochalasin D resulted in a significant increase in neuronal cell death in OHSCs that were challenged with Aβ1-42 oligomer-enriched preparations (Figure 5E). We thus
confirmed and elaborated recent findings that microglia in OHSCs take up Aβ (33, 91), which corroborates in vivo studies demonstrating that microglia from wild-type mice efficiently engulf Aβ (53, 83). By reducing the Aβ load in the brain, microglia are thought to contribute to neuropro-tection against AD pathology (21, 30, 31). Nevertheless, only a few studies have experimentally demonstrated the effects of microglial Aβ phagocytosis on neuronal damage (66, 84). By investigating Aβ plaque load and neuronal cell death following the depletion of microglial cells and by using microglial uptake blockers, the present study provides evidence showing that microglial Aβ engulfment is involved in the beneficial effects of microglia in OHSCs.
Our study demonstrates a potential protective role for functional ramified microglia against Aβ-mediated apoptotic mechanisms, most likely via clearance of Aβ deposits.
ACKNOWLEDGMENTS
This study was funded by the Competence Network Degenerative Dementias (KNDD) (project number 01GI1008C) and the Faber-Stiftung, Marburg.
CONFLICT OF INTEREST
The authors declare no conflict of interest concerning the content of the article.
DATA AVAILABILITY STATEMENT
The authors confirm that all data underlying the findings are fully available without restriction. All relevant data are within the paper.
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